A false coloured ESEM image of the epidermis of Vicia faba (Broad bean). The guard cells are turgid and the stomatal pores widely open.

Fig. 1: A false coloured ESEM image of the epidermis of Tradescantia andersonia. Three pores are visible, each formed by a pair of guard cells. The tissue was not subject to preparation by chemical or cryogenic preservation techniques and was not metal coated. Accelerating voltage 10 kV, beam current 0.09 nA, water vapour pressure 7.2 torr, 7°C.

Fig. 2: A sequence of three ESEM images following the closure of a single stomatal pore in response to the reduction in chamber relative humidity from 97% to 91%. (A) 12 minutes, (B) 25 and (C) 32 minutes after cutting. Accelerating voltage 10kV, beam current 0.09 nA. Water vapour pressure 7.3 torr, temperature 7°C in image (A), subsequent pressure 7.2 torr, temperature 8°C , in (B) and (C).

Historically, electron microscopy of dynamic biological processes has been impossible to achieve in real time because conventional electron microscopy requires specimen fixation, dehydration and metallic coating. The advent of the Environmental Scanning Electron Microscope (ESEM) removed these restrictions, allowing fully hydrated samples to be imaged in their native state. This raises the possibility of secondary electron imaging of dynamic biological processes.

Fig. 3: A false coloured ESEM image of Vicia faba leaves infected with the fungal pathogen Uromyces Vicia faba. Fungal spores can be seen erupting through the epidermal tissue. Image taken with a 1.5kV beam, FEI spotsize 5.

Well Suited to Biological Imaging

The ESEM differs from a Conventional Scanning Electron Microscope (CSEM) in that a differential pumping system maintains a pressure of gas (typically water vapour) in the specimen chamber whilst the gun remains at high vacuum. Ionizing collisions between electrons and these gas molecules create positive ions which drift down onto the sample tending to neutralize specimen charge. In this way it is possible to image insulating samples without the need for metallic coating. The presence of water vapour in the chamber also means that a high relative humidity can be maintained and therefore samples can be imaged in a hydrated state without the need for dehydration and fixation. These features suggest that ESEM could be well suited to biological imaging, offering high resolution topographic images with a remarkable depth of field, yet without the need for sample preparation or the artefacts it may introduce. Our work focuses on optimizing the ESEM for biological use and most recently [1] has involved exploring the possibility of imaging of biological systems undergoing natural morphological changes.

Visualizing Stomatal Closure

The closure of stomatal pores in leaf epidermal tissue was chosen as a test case as leaf tissue is fairly robust, readily available and stomatal movements occur on a timescale suitable for ESEM imaging. Stomatal pores are present in almost every species of terrestrial plant and control gaseous exchange with the atmosphere. A typical ESEM image of tradescantia epidermis, with widely open stomata pores, is shown in figure 1. Each pore is surrounded by two guard cells, which can swell and change shape, opening or closing the pore depending on their turgor pressure.

In nature, stomatal movements occur in response to environmental cues. In the microscope chamber the temperature and hence relative humidity can be varied with a view to inducing stomatal closure.

By optimizing the microscope to view stomatal movements in living Tradescantia andersonia (Spiderwort) leaf tissue, we were able to demonstrate that it is possible to follow a biological system undergoing dynamic morphological changes in real time. Considerations included minimizing beam damage and reconciling the need for an adequate physiological temperature and a low gas pressure favourable for imaging, with the thermodynamic constraints on achieving a high relative humidity. An imaging protocol was developed utilizing a custom pumpdown protocol developed by Cameron and Donald [2], for a specimen held at 7°C. The lower epidermis of Tradescantia andersonia was held at a water vapour pressure of 7.3 torr and imaged with a 10 kV, 0.09 nA beam, blanked between imaging scans. Stomatal closure could be triggered and observed in a controlled way by increasing the temperature and thus reducing the relative humidity. Figure 2 shows a series of three images taken over a 20 minute period, following the closure of a single pore in response to a reduction in relative humidity.

Epidermis

The Epidermis is in direct contact with the external environment. It contains many important adaptations which allow plants to survive & reproduce on land. We will observe the most general adaptations as well as some exotic ones. The functions of many types of epidermal cells are well known but there are some specialized cells with unknown functions.

The epidermis is important in both vegetative and reproductive organs. It is treated here in a broad sense as the superficial layer (or rarely layers) on all differentiated parts of the plant in the primary state of growth. During secondary growth the epidermis is often replaced by Periderm.

Many features of the epidermis can be seen in whole mounts at low magnification with the compound microscope.

Stomata should receive special attention. Sections may reveal Guard Cells cut in more than one plane.

Note whether guard cells are in the same plane as rest of epidermis, or if they are raised or sunken.

Subsidiary Cells may or may not be present. The arrangement of subsidiary cellsand guard cells can be used to identify plants

Observing leaf stomata

It is possible to observe the impression of leaf epidermis cells on white wood glue. The stomata and guard cells are easily visible. The regular shape of the stomata makes it an ideal specimen for practicing drawing.

Cut the glue into shape using scissors and observe it with the microscope. If the glue is still water soluble after drying, then do not immerse the glue into water. The contrast is low, it is necessary to close the condenser aperture diaphragm.

Method:

Evenly spread a drop of water soluble wood glue on the bottom side of a leaf (the stomata are located on the bottom side).

Wait several hours or overnight for the glue to dry.

Carefully peel off the glue. It has become transparent.

Use scissors to cut the glue into shape and observe under the microscope. The leaf epidermis cells have left an impression on the glue, which can be observed.

Key message

An automated process using a cascade classifier allowed the rapid assessment of the density and distribution of stomata on microphotographs from leaves of two oak species.

Abstract

Stomatal density is the number of stomata per unit area, an intensively studied trait, involved in the control of CO2 and H2O exchange between leaf and atmosphere. This trait is usually estimated by counting manually each stoma on a given surface (e.g., a microphotograph), usually repeating the procedure with images from different parts of the leaf. To improve this procedure, we tested the performance of a cascade classifier to automatically detect stomata on microphotographs from two oak species: Quercus afares Pomel and Quercus suber L. The two species are phylogenetically close with similar stomatal morphology, which allowed testing the reuse of the cascade classifier on stomata with similar shape. The results showed that a cascade classifier trained on only 100 sample views of stomata from Q. afares was able to rapidly detect stomata in Q. afares as well as in Q. suber with a very low number of false positives (5 %/1.9 %) and a small number of undetected stomata (14.8 %/0.74 %), when partial stomata near the edge of the microphotographs were ignored. The remaining undetected stomata were due to obstacles such as trichomes. As an example of further applications, we used the positions detected by the cascade classifier to assess the spatial distribution of stomata and group them on the leaf surface. To our knowledge this is the first time that a cascade classifier has been applied to plant microphotographs, and we were able to show that it can dramatically decrease the time needed to estimate stomatal density and spatial distribution.

Guard cells emit an alkali-induced, blue fluorescence upon excitation by ultraviolet radiation (emission maximum energy at 365 nm).Fluorescence emission of guard cells was brighter than that of the neighbouring epidermal cells in a number of wild and cultivated plants including conifers, but the relative fluorescence intensity and quality was species-dependent.

Immersing leaves of these plants in chloroform for 30 s (thereby removing epicuticular waxes) significantly reduced the intensity of the fluorescence emitted by guard cells. This indicates that guard cell fluorescence could be due to either an increased concentration of fluorescing compounds (probably wax-bound phenolics), or a thicker cuticular layer covering the guard cells.

Given that the alkali-induced blue fluorescence of the guard cells is a common characteristic of all plants examined, it could be used as a rapid and convenient method for in situ measurements of the number, distribution and size of stomatal complexes.

The proposed experimental procedure includes a single coating of the leaf surface by, or immersion of the whole leaf in, a 10% solution of KOH for 2 min, washing with distilled water, and direct observation of the leaf surface under the fluorescence microscope.

Fluorescence images were suitable for digital image analysis and methodology was developed for stomatal counting using Olea europaea as a model species.